Methods Index

Amaxa Nucleofection
Con A Coverslips
Formaldehyde Fixation
Freezing and Thawing
Growing S2 Cells
Hemacytometer
Making a Stable Cell Line Expressing Your Favorite Gene
Splitting Mammalian Tissue Culture Cells
Tissue Culture Facility

Amaxa Nucleofection

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1. Add 8mL growth medium to dish (not flask); warm 30 min in incubator.

2. Prepare nucleofection solution:

Let sit at room temperature.

3. Prepare the flask: wash with PBS and trypsinize. Ad 1 pipette full of medium, place cells in 15 mL conical tube and spin, 2 centrifuge, setting 1 for 10 min.

4. Remove as much media from above the pellet as possible. Add 2 ug DNA and 100 uL nucleofection solution. Using the p1000 (want large pipette tip) pipette to resuspend the cell pellet. Add this solution to the cuvette.

5. Put in Amaxa and run program X-001. use a transfer pipette to add media from the cuvette to the dish. Put dish in incubator.


Con A Coverslips

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Requirements:
Acid washed coverslips stored in 100% ethanol.
ConA-­‐0.5mg/ml.
Large petridishes to store the coverslips.

1. Flame the acid washed coverslips and cool for 10 sec. Place the coverslips on top of the large petridish. Usually make 9-­‐14 coverslips at a time to save time.

2. Thaw the conA from freezer and apply around 400uL of conA on the first two coverslips or how much ever is needed to cover the coverslip in conA.

3. Wait for 10 sec. Suck out the excess ConA and then tilt the coverslips to a slanted position and suck out the remaining from near the edges.

4. Repeat the same for all the coverslips. Cover the petridish with another lid of petridish and allow a small opening to dry. If possible, use a vacuum pump to dry it faster.
The process usually takes 15 minutes and drying takes half an hour.

Storage: In a petridish, place a clean kimwipe and place the coverslips on top with the conA surface facing up. Cover with the lid. 10 min before needed, UV sterilize them. Alternatively, we can UV sterilize all of them after drying and open the dishes only in aseptic conditions or inside hood.

Formaldehyde Fixation

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Fixative stock: 37% Formaldehyde: small bottle under chemical hood in prep lab.
TritonX-100: 10% stock, in hood in prep lab

To make 10 mL of fixative:

To a 15 mL conical plastic tube add:
1mL of 37% formaldehyde
0.5 mL of 10% triton
1 mL of 10x PBS
dH2O to 10 ml mark (8.5 mL)

Make in fume hood

To fix the cells:

  • Gently rinse cells 2X in warm PBS (remove medium, replace with PBS,
    remove PBS, replace with fresh PBS, remove again, replace). Leave
    cells in this PBS—until you add the fixative).
  • removing final PBS rinse and add about 1 mL of fixative to the cells; fix cells in hood for 10 mins.
  • Rinse cells by dunking each coverslip in 3 beakers of PBS-Tween-Azide, 10 X per beaker.
  • Place coverslip back in dish with fresh PBS-Tween-Azide
  • If you are not staining immediately, store cells in PBS-Tween-Azide in fridge.
  • Important things to remember

    Wear gloves and be very careful of the fixatives!!!
    Remember which side of the coverslip has the cells.
    Discard used fixative into the appropriate hazardous waste container in
    the chemical hood in the main lab.

    Paraformaldehyde fixation

    Fixative Stocks:
    32% Paraformaldehyde: bottle under chemical hood in main lab.
    Detergent: 10% triton X 100

    To make 10 mls of fixative:
    Paraformaldehyde: 1 mL stock
    10% triton: 0.5 ml
    10X PBS -/- 1 mL
    add water make 10 mL (8.5 mL)

    Use same method as Formaldehyde fixation.


    Freezing And Thawing

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    Freezing

    Use healthy cells which have been fed within 24 hours of freezing; use flasks that are nearly confluent.

    Trypsinize cells in usual manner; spin out cells in 15 mL tube for 5-10 minutes.

    Remove supernantant and resuspend pellet in 1 mL cell freezing medium.

    Quickly transfer cell mixture to sterile tube labeled with the cell name, passage number, date and initials of the person freezing the cells.

    Wrap tube(s) in a few layers of paper towels and place in the -80°C freezer in ISB 262A.

    Leave in the freezer for 24-48 hours, then remove cells and place in a cane in the liquid nitrogen tanks.

    Fill out the freezing log

    Be sure liquid nitrogen freezer does not run out of nitrogen!

    Cells will remain “viable” for at least a year, usually several.

    Thawing

    Pre-warm 10 mL of medium in a small flask in 37°C CO2 incubator.

    Remove the tube of cells that you wish to thaw from the liquid nitrogen tank.

    Rapidly bring to 37°C by holding in the water bath and swirling.

    As soon as cells are thawed, bring tube to hood, wipe tube with 70% ethanol.

    Using sterile technique, transfer cells to the flask of warm medium.

    Incubate in CO2 incubator overnight; next day change the medium, removing dead cells and DMSO. (Viable cells will have stuck to the flask.)

    Alternate procedure:

    Transfer thawed cells to tube containing 10 mL of warm medium

    Spin in clinical centrifuge to pellet cells

    Resuspend pellet into 5 mL of warm medium

    place cells in a flask and grow in incubator as usual.

    Freezing Medium

    Recipes here

    Make normal culture medium at 85% volume (ie medium components for 100 mL, but only bring the volume to 85 mL) AND with 20% serum.

    Freezing Medium: for one flask of cells, take 850 µL of freezing medium and add 150 µL of DMSO (sterile).

    Mix well and leave on ice. Make this fresh every time you freeze cells

    Freezing S2 cells

    S2 Cells Spin down a 3-4 day old flask.

    Resuspend in 4 mL of S2 medium with serum and antibiotic.

    Add 1mL DMSO (20% final).

    Make 1mL aliquots.

    Put in -80 in styrofoam for 1 day.

    Transfer to liquid nitrogen.


    Growing S2 Cells

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    1. Cells are in suspension -- they are not dead! To make conditioned media: spin cells to bottom of conical tube (be gentle!!). Save the supernatant in a labeled tube that has the date and is labeled “conditioned medium”. Store in fridge if there is extra.
    2. Grow cells at room temperature, no CO2. Passage every 3-4 days. Dilute cells 1:5 each split and use 20% conditioned media. Method:

    a. Put cells from flask in tube; spin; take off supernatant and save (this is conditioned medium).
    b. Resuspend pellet (gently) in 5 mls media. Take clean flask.
    c. Add to clean flask:

    3. To induce gene expression (i.e. tubulin) add copper sulfate to the media. Cells are Neo resistant. Induce day prior to imaging; to image put on previously prepared ConA coated coverslips. Image 30 min after plating on ConA
    To make ConA coverslips: Product is C-2631 from sigma
    Made it like this: 100mgs, want 0.5 mg/ml. Add 200 mls water, syringe filter. Store at -20 in aliquots.
    See my lab book Jan 2006, page 50.
    To make medium for growth of drosphila cells:


    Hermacytometer

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    Counting cells by the use of a hemacytometer is a convenient and practical method of determining cell numbers in the case that the Coulter counter is out-of-order temporarily. (It is not that bad.) The hemacytometer consists of two chambers, each of which is divided into nine 1.0 mm squares. A cover glass is supported 0.1 mm over these squares so that the total volume over each square is 1.0 mm x 0.1 mm or 0.1 mm3, or 10-4 cm3. Since 1 cm3 is approximately equivalent to 1 ml, the cell concentration per ml will be the average count per square x 104.

    Hemacytometer counts are subject to the following sources of error:

    With careful attention to detail, the overall error can be reduced to about 15%. It is assumed that the total volume in the chamber represents a random sample. This will not be a valid assumption unless the suspension consists of individual well-separated cells.
    Cell distribution in the hemacytometer chamber depends on the particle number, not particle mass. Thus, cell clumps will distribute in the same way as single cells and can distort the result. Unless 90% or more of the cells are free from contact with other cells, the count should be repeated with a new sample. A sample will not be representative if the cells are allowed to settle before a sample is taken. Always mix the cell suspension thoroughly before sampling.
    The cell suspension should be diluted so that each such square has between 20 - 50 cells (2-5 x 10 5 cells/ml). A total of 300 - 400 cells should be counted, since the counting error is approximated by the square root of the total count. A common convention is to count cells that touch the middle lines (of the triple lines) to the left and top of the square, but do not count cells similarly located to the right and bottom. Hemacytometer counts do not distinguish between living and dead cells. A number of stains are useful to make this distinction. Trypan blue among others (Erythrosin B, Nigrosin) can be used: the nuclei of damaged or dead cells take up the stain. If more than 20% of the nuclei are stained, the result is probably significant. Although the trypan stain distinction has been questioned, it is simple and gives a good approximation.

    Materials

  • 1. Clean hemacytometer and cover glass, or cover slips
  • 2. Pasteur Pipets or Transfer Pipets
  • 3. Balanced Salt Solution (BBS) or PBS
  • 4. Trypan blue, 0.4% in BBS (or PBS)
  • 5. Microscope
  • 6.Tubes
  • 7. Hand counter (Colony counter can be used)
  • 8. Cell suspension
  • Procedure

  • 1. Dilute 0.2 ml of Trypan blue with 0.8 ml of BBS.
  • 2. Place cover glass over hemacytometer chamber.
  • 3. Transfer 0.5 ml of agitated cell suspension to a 15 ml tube and add 0.5 ml of diluted trypan blue.
  • 4. With a Pasteur or transfer pipet, fill both chambers of the hemacytometer (without overflow) by capillary action. Cells will settle in the tube and in the pipet by gravity within a few seconds. Work quickly.
  • 5. Using the microscope with a 10X ocular (and a 10X objective), count the cells in each of 10 squares (1 mm2 each). If over 10% of the cells represent clumps, repeat entire sequence. If fewer than 200 or more than 500 cells are present in the 10 squares, repeat with a more suitable dilution factor.
  • 6. Calculate the number of cells per ml, and the total number of cells, in the original culture as follows: Cells/ml = average count per square x 104 Total cells = cells per ml X any dilution factor X total volume of cell preparation from which the sample was taken.
  • 7. Repeat count to check reproducibility (+/- 15%).

    References:

  • 1. Berkson, J., T. B. Magath and M. Hurn (1939). Am. J. Physiol. 128, 309.
  • 2. Sanford. K.K., W.R. Earle, V.J. Evans, H.K. Waltz and J.E. Shannon (1951).
  • 3. Absher, M. in Tissue Culture Methods and Applications, Eds. Kruse, P.F. and Patterson, M.K., Jr. Academic Press, N.Y., 1973, p.395.
  • From the Laboratory of Dr. Allan Bradley

    Baylor College of Medicine, Houston, Texas


    Making a Stable Cell Line Expressing Your Favorite Gene

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    1. Make a death curve. This is to determine the minimal concentration of antibiotic that kills cells that do not have the resistance gene.
    2. transfect the cells with the plasmid containing the gene that you want to express in the cells (see Amaxa protocol).
    3. incubate the cells for about 48 hours after transfection so that the gene is expressed, and the resistence gene is expressed.
    4. add the appropriate dose of antibiotic (this is the selection step). The cells that do not contain the resistance gene (i.e. that were not successfully transfected) will die. Change the medium every 2 or 3 days to remove dead cells. Replace medium with fresh medium containing the antibiotic.
    5. split the cells as needed. Use medium with antibiotic. Make some coverslips to see if the cells are expressing the gene of interest.

    After ~2 weeks any cell without the transgene and selectable marker should have died. The remaining cells are a **mixed population** – different cells might express different levels of the transgene. These can be used for preliminary experiments.

    **Making a clonal cell line**

  • A **Clonal cell line** contains cells that are derived from a single cell, and thus are a clone. They will all be genetically identical; this can be very helpful for experiments.
  • a. Trypsinize your 'mixed population' of cells; keep some growing in a flask
    and freeze some in case your cloning does not work the first time and you need to repeat the procedure.
  • b. take 50ul of the trypsinized cells, dilute again into 1 ml medium. Add 10 -150 ul of this dilute solution of cells to 100mm dishes containing growth medium. Look under the tissue culture scope. The cells should be very dilute – one or just a few cells per field of view. If they are too dense or dilute, adjust accordingly. You want each individual cell to land on the plastic dish and grow into a colony of cells that is separate from other colonies on the dish.
  • c. wait about 1 week for the colonies to grow.
  • d.isolate individual colonies. These are potential clonal cell lines.
  • Toss any cell lines that are not healthy or have too much or too little fluorescence.


    Splitting Mammalian Tissue Culture Cells

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  • 1. Cells are in suspension -- they are not dead! To make conditioned media: spin cells to bottom of conical tube (be gentle!!). Save the supernatant in a labeled tube that has the date and is labeled “conditioned medium”. Store in fridge if there is extra.
  • 2. Grow cells at room temperature, no CO2. Passage every 3-4 days. Dilute cells 1:5 each split and use 20% conditioned media. Method:

    a. Put cells from flask in tube; spin; take off supernatant and save
    (this is conditioned medium).

    b. Resuspend pellet (gently) in 5 mls media. Take clean flask.

    c. Add to clean flask:

    1 ml conditioned medium (supernatant)
    1 ml of cell suspension
    3 mls of fresh media

  • 3. to induce gene expression (i.e. tubulin) add copper sulfate to the media. Cells are Neo resistant. Induce day prior to imaging; to image put on previously prepared ConA coated coverslips. Image 30 min after plating on ConA.
  • To make ConA coverslips: Product is C-2631 from sigma

    Made it like this: 100mgs, want 0.5 mg/ml. Add 200 mls water, syringe filter. Store at -20 in aliquots.

    See my lab book Jan 2006, page 50.

    To make medium for growth of drosphila cells:


    Tissue Culture Facility

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    Hoods


    When beginning work in a hood, spray all walls and the working surface with 70% ethanol and wipe clean.
    When you have finished working in a hood: please leave it empty, spray the walls and working surface with 70% ethanol and wipe, and make sure that the gas is off.
    Leave the fan on and turn on the UV light.

    Incubators

    Immediately remove any contaminated dish or flask from the incubator and treat it with 30% bleach. Wipe the shelf it was on with 70% ethanol. Notify any individuals with cells in the incubator of the contamination.

    If you are using an incubator that requires CO2: Always check the water level in the bottom of the incubator, if level is low, add water. You are responsible for checking the level of CO2 in the tanks, if pressure falls below 800 psi the tank is emptying. After having received instructions, connect a full tank to the incubator and order a replacement (or notify someone).

    Microscopes

    Please make sure the microscopes are clean and covered when not in use.

    Pipets


    Dispose of all glass Pasteur pipets in the glass only bin.
    Disposable plastic pipets should be placed in trash (in the plastic sleeve).
    Reusable pipets should be placed tip up in the pipet jar.

    Supplies


    Label all items in the refrigerator with your name and date. Items that not labeled will
    be discarded.

    Vacuum Pump


    When you are done, please empty and rinse flask with 30% bleach.

    Other things to remember


    Keep the area clean at all times.
    Dishes and flasks of cells to be discarded should be treated with 30% bleach before
    trashing.